Proteomics is the large-scale study of proteins, particularly their structure and function. With the completion of the human genome project it has become clear that further inquiry into the interactions of proteins, both with other proteins and with nucleic acids, is necessary to understand how genomic data is processed. For example, the current understanding of the human genomic data gleaned to date suggests that there are approximately 33,000 coding regions in the human genome. However, approximately 200,000 different proteins have already been identified in humans. Determining how roughly 33,000 genes give rise to roughly 200,000 different proteins is a question central to proteomics.
Current methods to investigate protein functions are generally based on static analyses, rather than on dynamic observations. For example, technologies currently relied upon to investigate protein characteristics use mass spectroscopy, x-ray crystallography, nuclear magnetic resonance (NMR), and gel electrophoresis. By definition, these techniques investigate the structure of proteins based on static measurements such as size, charge, mass, magnetic moment, or a combination of those features. As a consequence, these analytical methods are not amenable to investigating peptides or proteins when their physical characteristics are in flux. Because protein interactions are dynamic and complex, these static approaches reveal only limited forms of data.
One feature of proteins that is inherently dynamic, their interactions with other molecules, is measured almost entirely via static methods. For example, ligand/receptor interactions are commonly visualized by their co-crystallization, which crystallizes the components at an arbitrarily fixed point in their interaction. Because the interaction of proteins involves changes in their mass, size, or charge, the process of protein-protein interaction is inferred from a comparison of the starting materials with the products. For example, the effectiveness of proteases on particular substrates is often inferred by incubating the putative protease with a putative substrate for some defined period of time. This is followed by subjecting the reaction mixture to SDS-PAGE gel separation to visualize the products. While this method of determining the effectiveness of a protease on a substrate is easily interpreted, it does not provide quantitative data, it is not continuous, and it is time consuming. Consequently, such methods are not amenable to high-throughput screening of protease activity specifically, or protein interactions generally.
Other methods of determining protease activity are equally limited. For example, enzyme-linked immunosorbent assays (ELISA) are flexible but are indirect, time-consuming, provide limited quantification, and are limited to discrete time points. Similarly, analytical methods that rely upon labeling free amino acid groups are limited to peptides. These methods are also time-consuming, and yield data that are limited to discrete time points. Other methods, such as measuring absorbance of light by a solution at 280 nm or measuring fluorescence intensity based on precipitation, are limited to non-specific proteases and are not continuous.
That being said, certain methods using fluorescence labeling allow for real-time monitoring of protein interactions. These known methods, however, suffer from other technological limitations. For example, conventional labeling of proteins with a fluorophore is not easily controlled. Multiple lysine or cysteine residues of the peptide being studied are usually labeled, thus resulting in potentially multiple fluorescent moieties being present in a single product. Consequently, the extent of fluorophore labeling becomes a variable that must be closely monitored and controlled. If the extent of labeling is uncontrolled or unknown, there will be multiple fluorescent moieties that will introduce unknown errors in the results (or might even confound the results entirely). Another known analytical method, fluorescence resonance energy transfer (FRET), requires extensive substrate optimization, is generally limited to peptides, and can also result in non-specific labeling. Thus, the fluorescence methods currently available require specific technical knowledge and specialized equipment for their optimal use. And, as in the other method discussed, the conventional fluorescent approaches are not well-suited for high-throughput screening.
While the conventional dynamic methods are cumbersome, fluorescence polarization yields valuable knowledge. Fluorescence polarization is based upon two physical phenomena. The first phenomenon is that a fluorescent molecule excited by plane-polarized energy emits energy in the same plane. The second phenomenon is that molecules in solution rotate naturally, with smaller molecules rotating more rapidly as compared to larger molecules. Because of these two phenomena, a correlation can be made between the size of a fluorescent molecule and the rate of change of its emitted, plane-polarized energy. Fluorescence polarization can be used to measure DNA-protein, protein-protein, and antigen-antibody interactions.
While the term “fluorescence polarization” is used herein, the term “fluorescence anisotropy” is often used synonymously to denote the same phenomena. Anisotropy is the characteristic of a medium wherein a given physical property differs in value in different directions within the medium. In the current instance, fluorescence anisotropy and fluorescence polarization are two different means to describe the same phenomenon. Anisotropy values and fluorescence values can be inter-converted using simple and well-known algebraic functions.
Recently a new method of fluorescently labeling a protein has been described by Griffin et al. (Science, 1998, 281: 269-272), Adams et al. (J. Am. Chem. Soc., 2002, 124: 6063-6076), and Tsien et al. (U.S. Pat. Nos. 5,932,474; 6,008,378; 6,054,271, and 6,451,569), all incorporated herein by reference. These documents describe biarsenical compounds and the specific target sequences to which they bind. An analytical method and corresponding kit utilizing these reagents is marketed under the trademark “FlAsH”-brand (Fluorescein Arsenical Hairpin binder) fluorescent labeling kit, and “ReAsH” (Resorufin Arsenical Hairpin binder), both by PanVera Corporation (Madison, Wis., now a wholly-owned subsidiary of Invitrogen Life Sciences, Carlsbad, Calif.). See Invitrogen Product Nos. P3050 for FlAsH, and P3006 for ReAsH, and product literature #L0920 Rev. 01/03).
This labeling method utilizes a reagent comprising a fluorescent ligand that is a biarsenical derivative of fluorescein and is weakly bound to a quenching moiety. The fluorescent ligand binds with greater affinity to a fluorescence tag, comprising a tetracysteine motif While neither the ligand nor the fluorescence tag is fluorescent alone, mixing them allows the fluorescence tag to displace the quenching moiety, thereby resulting in a highly fluorescent complex specifically bound to the tetracysteine motif. Because the motif is very rare in naturally occurring proteins, the fluorescence tag can be engineered into a protein with the expectation that the biarsenical ligand will bind specifically to the tetracysteine motif Currently, the FlAsH/ReAsH-branded systems are used for protein labeling investigations where the object is to follow a protein through in-vitro or in-vivo pathways.
As noted earlier, the progress to date of the human genome project indicates that the “one-gene, one protein” hypothesis will require considerable modification to account for the observed diversity of the proteome. The ability to use current techniques to further elucidate the role of proteins in their interactions and their role in developing protein diversity is extremely restricted by the limitations of current technology. Therefore, there is a long-felt and unmet need for a method to study protein interactions that allows for high-throughput screening of those interactions with a minimum of substrate optimization and product manipulation.